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Citation Palmer, Cody. 2017. PRO136ALA CTLA4 Mutation. Master's thesis,Harvard Medical School.

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PRO136ALA CTLA4 Mutation

CODY PALMER

A Thesis Submitted to the Faculty of

The Harvard Medical School

in Partial Fulfillment of the Requirements

for the Degree of Master of Medical Sciences in Immunology

Harvard University

Boston, Massachusetts.

May, 2017

ii

Thesis Advisor: Dr. Shiv Pillai Cody Palmer

PRO136ALA CTLA4 Mutation

Abstract

Cytotoxic T lymphocyte antigen–4 (CTLA-4) is an inhibitory receptor involved in the regulation of

immune responses. Deficiency of Ctla4 in mice causes fatal multiorgan lymphocytic infiltration. We identified a

heterozygous, missense c.406C>G, p.P136A mutation in CTLA4 in a CVID patient. This mutation is within the

binding motif of CTLA-4 and is predicted to interfere with ligand binding. To test whether this mutation is causal in

the phenotype of the patient, we will use CRISPR/Cas9-mediated genome editing to clone the mutation into a

regulatory T cell-like cell line and use the edited cell line for functional studies.

iii

Table of Contents 1. Chapter 1: Background .......................................................................................................................................1

1.1. Background ..................................................................................................................................................1

1.2 Schematic figures .........................................................................................................................................8

2. Chapter 2: Data and Methods ...........................................................................................................................10

2.1. Short Introduction .....................................................................................................................................10

2.2. Materials and Methods .............................................................................................................................10

2.3. Results ........................................................................................................................................................18

3. Chapter 3: Discussion and Perspectives ..............................................................................................................28

3.1. Limitations .................................................................................................................................................28

3.2. Future Research ........................................................................................................................................30

4. Bibliography .......................................................................................................................................................34

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Figures

Figure 1 CRISPR in bacteria and archaea

Figure 2 CRISPR in genome engineering

Figure 3 Schematic of workflow

Figure 4 Schematic of Cas9D10A

Figure 5 Targeting strategy and gRNA plasmid construction MLM3636

Figure 6 ssODN HDR template design

Figure 7 Nucleofection efficiency controls

Figure 8 Nucleofection results

Figure 9 Targeting strategy and gRNA plasmid construction pSpCas9n(BB)-2A-GFP

Figure 10 Sleeping Beauty transposon and transposase plasmids

Figure 11 ssODN HDR template design using CRISPR/Cas9-blocking mutations increase HDR accuracy by preventing re-editing by Cas9

v

Acknowledgements

I would like to express my sincerest gratitude to the Pillai lab at the Ragon Institute of MGH, MIT, and Harvard for all of the assistance and guidance they have provided during this work.

This work was conducted with support from Students in the Master of Medical Sciences in Immunology program of Harvard Medical School. The content is solely the responsibility of the authors and does not necessarily represent the official views of Harvard University and its affiliated academic health care centers.

1. Chapter 1: Background

1.1. Background

Systems of host defense have evolved since the first primitive immune system that was in existence by the

time that plants and animals diverged. Unicellular organisms such as bacteria have simple immune systems that

protect against bacteriophage infections. Other defense mechanisms evolved in eukaryotes. These mechanisms are

the basis of the innate immune system and include phagocytosis, antimicrobial peptides, and the complement

system. Jawed vertebrates and higher members of subphylum Vertebrata have evolved more complex defense

mechanisms that includes the ability to form immunological memory after an initial response to a specific pathogen

which allows for a faster and more efficient immune response upon re-exposure to that pathogen1.

Humans and other vertebrates of the class Mammalia have an adaptive immune system that consists of

lymphocytes that are able to recognize specificity among pathogens via antigen receptors on their surface that are

generated through somatic recombination. The adaptive immune system allows for more efficient and targeted

immune responses based on recognition of a large number of self- and non-self antigens. With this capacity to

generate such a large repertoire of immune cells comes the generation of self-reactive T cells and, thus, increased

opportunities for inappropriate responses against self-antigens1. Therefore, it is critical that mechanisms are in place

that aim to prevent these autoimmune events. Tolerance is controlled by multiple mechanisms, including inhibitory

receptors and regulatory T cells.

Immune system dysregulation can lead to autoimmune diseases, inflammatory diseases, and cancer.

Inflammation is a normal component of the biological response of tissue to harmful stimuli. The function of

inflammation is to eliminate the initial cause of injury and to initiate tissue repair. Inflammatory diseases are caused

by an inappropriate inflammatory response to an antigen that should not elicit an immune response under normal

circumstances. Cancer is also the result of a disordered immune response. The immune system normally is capable

of recognizing and eliminating cells that have undergone malignant transformation. The loss of this

immunosurveillance ability is pathogenic in the development of cancer. Immunodeficiency is the result of a

hypoactive immune system which causes recurring infections. Immunodeficiency can be the result of either a

genetic disease such as severe combined immunodeficiency, acquired conditions such as HIV/AIDS, or the use of

immunosuppressive medication. In contrast, autoimmunity is caused by a hyperactive immune system which forms

2

an immune response to self-tissues. Common autoimmune diseases include diabetes mellitus type 1, systemic lupus

erythematosus, and rheumatoid arthritis2, 3.

Although it may seem unlikely due to their seemingly opposite phenotypes, autoimmunity and

immunodeficiency can manifest concurrently in a single individual2. The mechanisms behind this association are not

fully known but is commonly seen in disorders such as CVID. Common variable immunodeficiency (CVID) is the

most common clinical primary immunodeficiency. The immune disorder is defined by high susceptibility to

infection, hypogammaglobulinemia, and impaired specific antibody response. By definition, all CVID patients have

low serum IgG with low IgA or low IgM, and impaired antibody response to vaccination. The symptoms of CVID

are highly variable, as the name suggests. Most commonly, patients have recurrent bacterial infections, usually of

the upper respiratory tract. Infections are a direct result of the low antibody levels in the circulation, which do not

adequately protect against pathogens. The most frequent cause of infections in CVID are due to the bacteria

Haemophilus influenzae and Streptococcus pneumoniae. Infections mostly affect the sinopulmonary tract, however,

they can also occur at other sites, such as the eyes, skin and gastrointestinal tract. Pneumonia, bronchitis, and

sinusitis are common in people with CVID. About 50% of patients have at least one occurrence of pneumonia.

These infections can become recurrent, leading to chronic lung disease. This chronic lung diseases manifests as

bronchiectasis or interstitial lung disease. Patients with interstitial lung diseases are most likely to also have

autoimmunity. Recurrent and chronic diarrhea is seen in approximately 40% of patients, frequently due to Giardia,

Salmonella and Campylobacter jejuni. Acute and chronic gastritis related to H. pylori infection is also seen.

Over 50% of patients also develop complications, including enteropathy, lymphocytic infiltration of tissues,

malignancy, and autoimmunity. Immune thrombocytopenic purpura and autoimmune hemolytic anemia are the most

common autoimmune features, but other manifestations include rheumatoid arthritis, inflammatory bowel disease

(IBD), systemic lupus erythematosus, diabetes mellitus, multiple sclerosis, and psoriasis. Accordingly, CVID

presents in many patients as an immune dysregulation syndrome with variable symptoms. CVID has a nonspecific

clinical presentation and laboratory definition which makes diagnosis difficult. Diagnosis of CVID is based mostly

on exclusion of other causes, such as secondary immunodeficiencies. The cause of CVID is unknown in

approximately 90% of cases. An autosomal dominant inheritance pattern is observed in most cases of familiar

CVID. Most mutations that are causal to CVID or increase susceptibility to diseases have not been identified3, 4, 8.

3

Most cases of CVID are sporadic and occur in people with no apparent family history. Not all individuals

who inherit a gene mutation will develop CVID. In many instances, affected children have an unaffected parent with

the same mutation. Penetrance may also appear incomplete due to late onset of symptoms. It is likely that both

environmental and genetic factors play causal roles. Although the specific environmental factors are unclear,

mutations in at least 13 genes have been associated with CVID. However, these known mutations only account for

approximately fifteen percent of cases. Since CVID is characterized by defective differentiation of B cells to plasma

cells and memory B cells, it is believed that the genetic influences in CVID are mutations in genes that are involved

in the development and function of B cells. Mutations in genes that encode BCR-associated complex or are involved

in BCR signaling (CD19, CD81, CD21) or are involved in B cell development and plasma cell differentiation

(CD20) have been associated with CVID. The most frequent mutations occur in the TNFRSF13B gene, also known

as TACI, a receptor for BAFF and APRIL. The protein produced from this gene plays a role in the survival and

maturation of B cells and in immunoglobulin production. TNFRSF13C, a receptor for BAFF also known as BAFFR,

is also associated with CVID. Other mutations associated with CVID are in genes involved in the function and

maturation of immune cells, such as T cells. It has been observed that, in addition to intrinsic B cell defects, CVID

can be caused by deficient T cell co-stimulation. Accordingly, alterations in the function and frequency of T cells

have also been documented in patients with CVID. Mutations in the T cell co-stimulatory molecule ICOS as well as

the T cell inhibitory molecule CTLA-4 have been identified in CVID. These defects may cause changes in the

interaction between B and T cells which may provide potential mechanisms into the various complications seen in

CVID3, 5, 6.

There are different types of CVID that are distinguished by differing genetic causes. Patients with the same

type of CVID may have varying symptoms and clinical presentation. The heterogeneity of CVID has led to several

attempts to classify CVID into subgroups based on clinical presentation and immunological features. When a

molecular diagnosis is made, the disease becomes a specific genetic deficiency and is considered distinct entities

separate from CVID. However, it is likely that CVID is a common disease that can result from many various genetic

mutations and in the future, these subcategories will be re-categorized as CVID with specific molecular etiology that

will not affect classification of the disease but rather give physicians a molecular basis for personalized treatment,

much like the way cancer is treated.

4

CTLA4 mutations were first reported in subjects clinically diagnosed with CVID. Heterozygous Ctla4

deficiency in mice shows no apparent phenotype. However, heterozygous human CTLA4 deficiency causes immune

dysregulation characterized by recurrent infections, hypogammaglobulinemia, and various autoimmune

manifestations, much like symptoms characteristic of CVID. However, instead of falling under the classification of a

complex disease with multifactorial causes like most cases of CVID, this phenotype has a monogenic cause and can

be treated accordingly7, 8.

The inhibitory receptor CTLA-4 is an excellent therapeutic target. As a key molecule in the regulation of

the immune response, CTLA-4 expression can be therapeutically modulated in order to reduce or increase immune

activity. CTLA-4 is a critical component of the effector function of regulatory T (Treg) cells. Treg cells are used by

the mammalian immune system as a mechanism of tolerance to control self-reactive T cells. Treg cells are a

subpopulation of T cells which control immune homeostasis and tolerance to self-antigens. This population of cells

are immunosuppressive and function to downregulate the proliferation of effector T cells. Naturally occurring

CD25+CD4+ Treg cells specifically express Forkhead box P3-positive (FOXP3+) which is a master regulator of Treg

cell development and function. Mutations in FOXP3 resulting in deficiency of Treg cells cause an autoimmune

syndrome called IPEX (immune dysregulation polyendocrinopathy X-linked). In mice, homozygous deficiency of

Ctla4 results in a fatal immune dysregulation with clear similarities to FOXP3 deficiency9-12. CTLA-4 is

constitutively expressed by Treg cells and mediates their suppressive function through the inhibition of naive T cell

activation. CTLA-4 is also expressed by T cells upon activation. Upon ligation to ligand, CTLA-4 inhibits

proliferation of activated T cells1. The modulation of inhibitory immune receptors such as CTLA-4 is changing the

way many conditions and diseases are being treated including inflammatory diseases, immunodeficiencies, and

cancer.

CTLA-4 was only recently discovered in 1987 when scientists at the Centre d’Immunologie in France

isolated cDNA clones from activated CD8+ T cells that defined a sequence encoding a 223 amino acid protein which

they called cytotoxic T cell antigen-4 (CTLA-4). Structural features of CTLA-4 identified the protein as a member

of the immunoglobulin superfamily. Subsequent to the identification of CTLA-4, it was cloned and mapped to the

same chromosomal region in both human and murine genomes as another member of the immunoglobulin

superfamily, CD28. CTLA-4 is homologous to the T cell co-stimulatory protein CD28. CTLA-4 was identified as a

second receptor for the T cell co-stimulation ligand B7 in 1991. The genetic and molecular similarities between

5

CTLA-4 and CD28 suggested that the two T cell surface molecules may also share similar functionality. However,

the function of CTLA-4 remained largely unknown until 1995 when it was identified as an inhibitor of T cell

activation13-15.

In order to become properly activated, T cells requires two signals. The first signal is provided by binding

of the T cell receptor (TCR) to its cognate peptide presented on major histocompatibility complex (MHC) on an

antigen-presenting cell (APC). This signal provides specificity to the subsequent immune response based upon the

identity of the present antigen. The second signal comes from co-stimulation, in which B7 co-stimulatory molecules

on the APC bind to the surface receptor CD28 on the T cell. Naive T cells express the co-stimulatory receptor

CD28, so co-stimulation for these cells comes from the CD80 and CD86 proteins, which together constitute the B7

protein, (B7-1 and B7-2, respectively) on the APC. The expression of B7 is induced by various stimuli, such as

products of pathogens or cell damage, in order to restrict immune activation only when appropriate. Co-stimulation

is requisite to appropriately activate T cell functionality. In the absence of co-stimulation, TCR signaling results in

anergy. On re-stimulation, anergic T cells are unable to produce IL-2 or to proliferate, even in the presence of co-

stimulatory signals. This is one of the mechanisms the immune system uses to ensure that T cells are only activated

in appropriate circumstances. CD28 co-stimulation initiates intracellular signaling that increases cellular

metabolism, proliferation, and differentiation16, 17.

After the immune system is activated, the strength and duration of immune responses must be tightly

regulated. The degree of T cell activation is determined by ligation of activating versus inhibitory receptors of the

CD28 gene family. Upon activation of the T cell, other receptors are expressed, such as ICOS and PD-1, that

provide additional co-stimulatory and inhibitory signaling. T cell activation through the TCR and CD28 also leads to

increased expression of CTLA-4. CTLA-4 is constitutively expressed in Treg cells but only upregulated in

conventional T cells after activation. CTLA-4 is homologous to CD28 and both molecules bind to B7-1 and B7-2 on

APCs. B7-1 and B7-2 are shared ligands of CTLA-4. The expression profiles of B7-1 and B7-2 differ. B7-2 is

constitutively expressed in APCs at low levels and rapidly up-regulated upon activation, whereas B7-1 is expressed

only upon activation. CTLA-4 binds mostly to B7-1 while CD28 is the preferential receptor for B7-217-22. Upon

ligation, CTLA-4 downregulates immune responses and prevents the over activation of T cells by inhibiting T cell

proliferation, differentiation, cell cycle progression, and IL-2 production. It has recently been shown that CTLA-4

functions by binding to and physically removing B7 ligands from APCs via a process called transendocytosis.

6

Through this ligand downregulation, CTLA-4 reduces activation of conventional T cells by reducing the availability

of co-stimulatory ligands on APCs, thus suppressing T cell activation24.

CD28 and CTLA-4 both bind B7 via the same motif. A hexapeptide motif (99MYPPPY104) in the FG loop,

conserved between CD28 and CTLA4, is essential for ligand binding. Mutagenesis experiments have shown that this

sequence is essential for binding to B7 and alanine substitutions of these residues reduce or abolish binding.

Although CD28 and CTLA-4 share B7 as ligands, CTLA-4 binds with greater avidity and affinity thus enabling it to

outcompete CD28 for its ligands. CTLA-4 is able to outcompete CD28 for B7 due to the manner in which it binds.

CTLA-4 is able to bind B7 bivalently, resulting in a very stable lattice structure of alternating CTLA-4 and B7

dimers. This oligomeric binding pattern displaces CD28 from the synapse due to steric crowding18-23.

CD28 and CTLA-4 are closely linked on human chromosome 2q33. The structure of CTLA4 contains four

exons, two untranslated regions upstream of exon 1 (5′ UTR) and downstream of exon 4 (3′ UTR), and a promotor

region up to −335 bp. Exon 1 encodes the signal peptide sequence; exon 2, an extracellular IgV-like domain that

contains the B7 binding domain; exon 3, the transmembrane region, and exon 4, the cytoplasmic tail.

Polymorphisms of the CTLA4 gene region have been associated with increased susceptibility to several autoimmune

diseases such as type 1 diabetes, celiac disease, rheumatoid arthritis, and multiple sclerosis12, 17. However, further

research is required to determine how polymorphisms directly affect CTLA-4 function.

Gene editing technology offers a powerful tool for modeling disease-associated mutations. Bacterial

CRISPR/Cas adaptive immune systems have been modified for use in editing of the eukaryotic genome. The

CRISPR/Cas system is a microbial immune system used by bacteria and archaea that provides acquired immunity

against viruses and plasmids by conferring resistance to foreign genetic elements (Figure 1). Type II CRISPR

systems incorporate foreign DNA within the host genome at CRISPR loci and the corresponding CRISPR RNAs

(crRNAs) are used to guide targeted degradation of homologous sequences. The recognition of exogenous DNA by

a Cas complex prompts the capture of viral or plasmid DNA and insertion of the exogenous DNA into the CRISPR

locus in the form of a novel repeat-spacer. Transcripts from the repeat-spacer array are processed into CRISPR

RNAs (crRNAs) that contain a sequence transcribed from the exogenous DNA called a protospacer sequence. The

part of the crRNA encoded by the protospacer directs Cas to cleave the complementary DNA target sequence if it is

adjacent to a short sequence called a protospacer adjacent motif (PAM) that is rarely present in the host genome, the

identity of which varies depending on the specific CRISPR system. The PAM sequence is used as a mechanism of

7

tolerance to prevent cleavage of host DNA and is requisite for binding of the Cas complex to the target sequence.

Transactivating CRISPR RNA (tracrRNA) binds to the crRNA and forms an active complex with Cas nuclease. The

active Cas complex uses the spacer target sequence to recognize and cut exogenous genetic elements29, 30.

CRISPR/Cas9 genome editing most commonly uses a Type II CRISPR system from Streptococcus

pyogenes. This system utilizes Cas9, crRNA, tracrRNA, and an optional repair template (Figure 2). Cas9, crRNA,

tracrRNA, and a repair template must be introduced into the target cell. The crRNA and tracrRNA can be joined

together to form a single-guide RNA (sgRNA). The sgRNA consists of the guide or target sequence and a scaffold.

The protospacer portion of the crRNA contains the target sequence that Cas9 uses to identify and directly bind to the

DNA of the cell. The target sequence is a 20-nt guide sequence that provides the specificity of the Cas9 nuclease.

When the Cas9 protein is expressed, it will form a complex with the sgRNA by binding to the scaffold domain of

the sgRNA. Cas9 is able to select the correct location on the host DNA by using the guide sequence to bond with

base pairs on the host DNA. For the S. pyogenes system, the target sequence must immediately precede a 5′-NGG

PAM sequence. The 20-nt guide sequence base pairs with the opposite DNA strand to mediate Cas9 cleavage 3 bp

upstream of the PAM sequence. Cas9 has two endonuclease domains, RuvC and HNH, which allows cleavage of

both strands of the target DNA resulting in a double-stranded break (DSB). DSBs are typically repaired by non-

homologous end-joining (NHEJ), which results in nonspecific insertions, deletions or other mutations. This is a

commonly used approach to engineer cell lines or animal models with specific gene knockout mutations. Specific

sequences can be inserted or replaced by using a DNA repair template to direct repair of DSBs by homology-

directed repair (HDR). A repair template introduced into the cell can be used as an alternative template in order to

produce specific nucleotide changes. The cell can use the repair template containing the chosen edit to repair the

break, thus incorporating the mutation of interest into the genome of the cell. Although CRISPR/Cas9 is widely used

to engineer gene knock-outs through NHEJ, the inefficiency of HDR prevents its widespread use for modeling

genetic disorders through the introduction of disease-associated mutations30-35.

8

1.2. Schematic figures

9

10

2. Chapter 2: Data and Methods

2.1. Short Introduction

Using a combined whole exome sequencing (WES) and targeted screening approach, we identified a novel

heterozygous, de novo, missense c.406C>G, p.P136A mutation in CTLA4 in a subject with CVID. Proline 136 is

within the binding motif of CTLA-4. A missense mutation within this ligand-binding sequence is predicted to result

in CTLA-4 deficiency due to an inability of the protein from the mutated allele to bind to B7. The protein product of

the allele carrying the mutation likely acts antagonistically to the wild-type allele. Since CTLA-4 functions as a

homodimer, the protein carrying the p.P136A mutation may dimerize with wild-type proteins forming a dimer that is

unable to oligomerize due to the inability of the mutant polypeptide to bind ligand. It is predicted that this mutation

acts in a dominant-negative manner, reducing the activity of the CTLA-4/B7 complex. It is unknown whether the

mutant protein is expressed at a normal level. If overexpressed, the proportion of CTLA-4 dimers containing only

normal subunits will decrease.

The clinical presentation of the patient was typical of CVID with autoimmune manifestations. Patient

history showed low IgG and IgA, low switched memory B cells, recurrent infections of the respiratory tract,

infiltrative granulomatous disease, ITP, and AIHA. To determine whether the CTLA4P136A mutation is causal in the

phenotype of the patient, we used CRISPR/Cas9-mediated genome editing to clone the mutation into a regulatory T

cell-like cell line for use in functional studies.

2.2. Materials and Methods

MLM3636 gRNA plasmid construction. The target sequence for the gRNA was selected based upon proximity the

intended Cas9 cleavage site with the lowest number of off-target sites. gRNA efficiency and off-target sites were

analyzed using a CRISPR design tool (http://crispr.mit.edu). gRNA sequences targeting CTLA4 were cloned into

plasmid MLM3636. MLM3636 was a gift from K. Joung (Addgene plasmid number 43860). Two DNA

oligonucleotides with vector-specific overhangs were ordered and then annealed. The top (FORWARD) and bottom

(REVERSE) phosphorylated oligonucleotides were suspended to 100 µM in 0.1x TE buffer and diluted 1:10 to 10

µM in H2O. The phosphorylated oligonucleotides were annealed in the reaction buffer detailed below.

Top strand oligo FOR (10 µM) 1 µl

Bottom strand oligo REV (10 µM) 1 µl

11

10x T4 DNA Ligase buffer 2 µl

dH2O (nuclease free) 16 µl

20 µl total volume

The annealing reaction was heated to 95°C for 5 mins and then cooled to 10°C. The double-stranded DNA product

was ligated into the BsmBI sites of the MLM3636 plasmid backbone. The ligation reaction is detailed below.

Plasmid backbone (cut MLM3636 vector) 1 µl

Annealed primers 1 µl

10x buffer (T4 DNA ligase buffer) 1 µl

T4 DNA ligase 0.5 µl

dH2O (nuclease free) 6.5 µl

10 µl total volume

The ligation reaction was incubated at 16°C overnight. The ligation product was transformed into E. coli. Bacterial

cells were thawed on ice. 1 µl of DNA in H20 and 25 µl of cells were combined and left on ice for 15 mins and then

heat shocked at 42°C for 45 s. The reaction volume was then placed back on ice for 2 mins. 250 µl of LB medium

was added to the reaction volume and was incubated at 37°C for 1 h. 100 µl of the reaction volume was plated on

each of four LB plates with antibiotics and left to grow at 37°C overnight.

The plates were inspected for E. coli growth and eight colonies were selected. Each colony was inoculated into a

loose-capped culture tube containing 5 ml LB medium plus antibiotic and incubated with 250 rpm shaking at 37°C

overnight. After 12-16 h of growth, the bacterial cells were harvested by centrifugation at 6800 x g for 20 mins.

Glycerol stocks of the eight colonies were prepared and stored at -80°C.

The plasmid DNA was isolated from the bacteria via miniprep. The pelleted bacterial cells were resuspended in 250

µl 2–8°C Buffer P1 with RNase A solution and transferred to a 1.5 ml tube. 250 µl Buffer P2 was added to the

bacterial cell solution and the tube was inverted 5 times to homogenize the solution. 350 µl Buffer N3 was added to

the solution and the tube was inverted 5 times to homogenize the solution. The solution was centrifuged in a

microcentrifuge at 13,000 rpm for 10 mins. The supernatant was decanted into a spin column. The spin column was

centrifuged at 13,000 rpm for 30 s and the flow-through was discarded. The spin column was washed with 0.5 ml

Buffer PB. The spin column was centrifuged at 13,000 rpm for 30 s and the flow-through was discarded. The spin

column was washed with 0.75 ml Buffer PE. The spin column was centrifuged at 13,000 rpm for 30 s and the flow-

through was discarded. The spin column was centrifuged at 13,000 rpm for 1 min to remove wash buffer. Plasmid

DNA was eluted from the spin column into a 1.5 ml tube by the addition of 50 µl Buffer EB (10 mM TrisCl, pH 8.5)

12

to the spin column. After 1 min, the spin column and 1.5 ml tube was centrifuged at 13,000 rpm for 1 min. DNA

concentration was determined by UV spectrophotometry at 260 nm. The sequences of the resulting clones were

verified via Sanger sequencing.

Bacteria from sample 1 and sample 2 were plated from glycerol stock on agar plates. The plates were incubated at

37°C overnight. Each colony was inoculated in 1–5 ml LB medium with antibiotic to form a starter culture and

incubated 4–6 hours at 37°C. Each starter culture was diluted into 150 ml LB medium and incubated for 12–16 h at

37°C at 300 rpm until cell density reached approximately 3–4 x 109 cells per ml. The bacterial cells were pelleted by

centrifugation at 6800 x g for 20 mins. Plasmid DNA was purified from the bacteria via maxiprep. The pelleted

bacterial cells were resuspended in 10 ml 2–8°C Buffer P1 with RNase A solution. 10 ml Buffer P2 was added to the

bacterial cell solution and the tube was inverted 5 times to homogenize the solution. The lysis reaction solution was

incubated at 25°C for 5 mins. 10 ml of pre-chilled Buffer P3 was added to the lysate and the tube was inverted 5

times to homogenize the solution. The lysate was transferred into a filter cartridge and the solution was incubated at

25°C for 10 mins. The HiSpeed maxi tip was equilibrated with Buffer QBT. 10 ml Buffer QBT was added to the

maxi tip and allowed to drain. After the precipitate of protein, genomic DNA, and detergent floated to the surface of

the lysate solution, a plunger was used to push the cell lysate through the filter cartridge into the equilibrated tip.

The tip was washed with 60 ml Buffer QC. The DNA was eluted from the tip filter with 15 ml Buffer QF. The DNA

was precipitated by adding 10.5 ml 25°C isopropanol to the eluted DNA. The solution was incubated at 25°C for 5

mins. The eluate/isopropanol solution was transferred into a 30 ml syringe with attached QIAprecipitator. The

eluate/isopropanol solution was filtered through the QIAprecipitator using a plunger. The QIAprecipitator was

removed and the plunger was removed from the syringe. The The QIAprecipitator was reattached to the syringe. The

DNA was washed by pressing 2 ml 70% ethanol through the QIAprecipitator using the plunger of the syringe. The

QIAprecipitator was removed and the plunger was removed from the syringe. The QIAprecipitator was reattached to

the syringe. The filter membrane of the QIAprecipitator was dried by using the plunger to press air through the

QIAprecipitator. This step was repeated. The QIAprecipitator was then attached to a 5 ml syringe. The DNA was

eluted from the filter membrane into a 1.5 ml tube using a plunger to press 1 ml Buffer TE through the syringe. The

QIAprecipitator was removed from the syringe, the plunger was removed, and the QIAprecipitator was reattached to

the syringe. The eluate was transferred from the 1.5 ml tube into the 5 ml syringe. The DNA was eluted again. DNA

concentration was determined by UV spectrophotometry at 260 nm.

13

HDR template construction. The 101-nt ssODN repair template was designed with homologous genomic flanking

sequence centered around the Cas9 cleavage site as described in Figure 6.

pSpCas9n(BB)-2A-GFP gRNA/Cas9 plasmid construction. The target sequence was the same guide sequence

selected for MLM3636. gRNA sequences targeting CTLA4 were cloned into plasmid pSpCas9n(BB)-2A-GFP.

pSpCas9n(BB)-2A-GFP (PX461) was a gift from Feng Zhang (Addgene plasmid # 48140). Two DNA

oligonucleotides with vector-specific overhangs were ordered and then annealed. The top (FORWARD) and bottom

(REVERSE) oligonucleotides were suspended to 100 µM in 0.1x TE buffer and diluted 1:10 to 10 µM in H2O. The

oligonucleotides were phosphorylated and annealed in the reaction buffer detailed below.

Top strand oligo FOR (10 µM) 1 µl

Bottom strand oligo REV (10 µM) 1 µl

10x T4 DNA Ligase buffer 1 µl

T4 Polynucleotide Kinase 1 µl

dH2O (nuclease free) 6 µl

10 µl total volume

The reaction was incubated at 37°C overnight to allow phosphorylation of the oligonucleotides to allow subsequent

ligation. The phosphorylated reaction mixture was then heated to 95°C for 5 mins and then cooled to 10°C. The

pSpCas9n(BB)-2A-GFP vector was digested in the reaction volume detailed below.

pSpCas9n(BB)-2A-GFP plasmid 3 µl

10x buffer (NEBuffer 2.1) 3 µl

BbsI (restriction enzyme) 2 µl

dH2O (nuclease free) 22 µl

30 µl total volume

The reaction was incubated at 37°C overnight to allow digestion of the vector backbone. The resulting concentration

of the vector was 75 ng/µl. The vector backbone was dephosphorylated according to the reaction outlined below.

Plasmid backbone (cut pSpCas9n(BB)-2A-GFP vector) 30 µl

Antarctic Phosphatase (AnP) 3 µl

10x buffer (AnP buffer) 4 µl

dH2O (nuclease free) 3 µl

40 µl total volume

The reaction was incubated at 37°C for 2.5 h. The reaction mixture was then heated to 80°C for 30 mins to heat

inactivate BbsI and AnP. The annealed oligo duplex was ligated into the BbsI sites of the pSpCas9n(BB)-2A-GFP

plasmid backbone. The ligation reaction is detailed below.

14

Plasmid backbone (cut pSpCas9n(BB)-2A-GFP vector) 0.5 µl

Annealed primers 1 µl

10x buffer (T4 DNA ligase buffer) 1 µl

T4 DNA ligase 0.5 µl

dH2O (nuclease free) 7 µl

10 µl total volume

The ligation reaction was incubated at 16°C overnight. The ligation product was transformed into E. coli. Bacterial

cells were thawed on ice. 10 µl of DNA in H2O and 100 µl of cells were combined and left on ice for 30 mins and

then heat shocked at 42°C for 45 s. The reaction volume was then placed back on ice for 2 mins. 250 µl of LB

medium was added to the reaction volume and was incubated at 37°C for 1 h. 150 µl, 100 µl, 60 µl, and 50 µl of the

reaction volume was plated on each of four LB plates with antibiotics and left to grow at 37°C overnight.

The plates were inspected for E. coli growth and 4 colonies were selected. Each colony was inoculated into a loose-

capped culture tube containing 5 ml LB medium plus antibiotic and incubated with 250 rpm shaking at 37°C

overnight. After 12–16 h of growth, the bacterial cells were pelleted by centrifugation at 6800 x g for 20 mins.

The plasmid DNA was isolated from the bacteria via miniprep. The pelleted bacterial cells were resuspended in 250

µl 2–8°C Buffer P1 with RNase A solution and transferred to a 1.5 ml tube. 250 µl Buffer P2 was added to the

bacterial cell solution and the tube was inverted 5 times to homogenize the solution. 350 µl Buffer N3 was added to

the solution and the tube was inverted 5 times to homogenize the solution. The solution was centrifuged in a

microcentrifuge at 13,000 rpm for 10 mins. The supernatant was decanted into a spin column. The spin column was

centrifuged at 13,000 rpm for 30 s and the flow-through was discarded. The spin column was washed with 0.5 ml

Buffer PB. The spin column was centrifuged at 13,000 rpm for 30 s and the flow-through was discarded. The spin

column was washed with 0.75 ml Buffer PE. The spin column was centrifuged at 13,000 rpm for 30 s and the flow-

through was discarded. The spin column was centrifuged at 13,000 rpm for 1 min to remove wash buffer. Plasmid

DNA was eluted from the spin column into a 1.5 ml tube by the addition of 50 µl Buffer EB (10 mM TrisCl, pH 8.5)

to the spin column. After 1 min, the spin column and 1.5 ml tube was centrifuged at 13,000 rpm for 1 min. DNA

concentration was determined by UV spectrophotometry at 260 nm. The sequences of the resulting clones were

verified via Sanger sequencing.

Sanger sequencing. 10 µl of plasmid DNA at a concentration of 200 ng/µl and 10 µl of sequencing primer at a

concentration of 1.5–3.0 µM were added to a 1.5 ml tube for submission. Human U6 promoter, forward primer was

15

used for sequencing.

MLM3636/Cas9D10A-GFP nucleofection. 106 MT-2 cells were nucleofected with 0.4 µg gRNA (MLM3636) and

1.6 µg Cas9 (Cas9D10A-GFP) in a 1:1 molar ratio with 1 µl ssODN template per 500 ng of plasmid DNA. Cells

used for nucleofection have >95% viability. 106 MT-2 cells were put into a 15 ml conical and washed once with

PBS with the supernatant aspirated out. The cells were resuspended in 100 µl of supplemented SE Nucleofector

solution (2.25 ml SE Nucleofector solution + 0.5 ml supplement). 2 µg of plasmid DNA and 1 µl of the ssODN

template was added to the 100 µl of cells in SE solution. 100 µl of the cell and DNA mixture was transferred into a

Nucleocuvette vessel and the vessel was placed in the 4-D Nucleofector X unit. The Jurkat CL-120 program was

selected and run. After the run (~1 second), the Nucleocuvette vessel was removed and the contents were transferred

in 2 ml of 37°C RPMI 1640 + 2 mM Glutamine + 10% Fetal Bovine Serum (FBS) + 5 ml P/S.

Live/dead viability staining. Cells were centrifuged at 1600 rpm for 5 mins and resuspended in 1 ml PBS. 1 µl of

live/dead viability dye was added. Cells were incubated with dye for 30 mins and centrifuged at 1600 rpm for 5

mins. Cells were washed with 1 ml PBS and centrifuged at 1600 rpm for 5 mins. Cells were resuspended in 200 µl

PBS.

Fluorescence-activated cell sorting. GFP+ cells were isolated and collected by FACS 48 h post-transfection and

resuspended in 50% conditioned RPMI 1640 + 2 mM Glutamine + 20% Fetal Bovine Serum (FBS) + 5 ml P/S.

Cell culture. MT-2 lymphocyte cell cultures were maintained at a cell concentration between 5 X 104 and 4 X 105

cells/ml in RPMI 1640 + 2 mM Glutamine + 10% Fetal Bovine Serum (FBS) + 5 ml Penicillin Streptomycin (P/S)

at 37°C 5% CO2.

Cell cloning by limiting dilution. Cells were homogenized by passing several times through a serological pipet. A

dilute cell solution (<106 cells/ml) was obtained before counting the cells in order to increase accuracy of the

number of cells present. The cell concentration in the homogenized cell solution was quantitated via cell counter.

Cells from the homogenized cell solution were transferred into the conditioned medium to make a new cell solution

at concentrations of 5 cells/ml, 10 cells/ml, and 50 cells/ml. 10 ml of the new cell solution at each concentration was

prepared per 96-well plate. 100 µl of the new cell solution was pipetted into each well of a 96-well plate. The 96-

well plates were placed in a 37°C 5% CO2 incubator for 7 days. The growth was observed 7 days and 10 days after

plating.

16

Restriction digest. Cas9D10A-GFP was digested using three enzymes as detailed below.

Buffer Enzyme Cuts Temperature

NEB Cutsmart XhoI 5289-5288 (9271 bp) (Linearize) 37°C

NEBuffer 3.1 BglII 1996-2932 (937 bp), 2933-1995 (8334 bp) 37°C

NEB Cutsmart EcoRI-HF 1728-5567 (3840 bp), 5568-1727 (5431 bp) 37°C

For each enzyme digest reaction, the enzyme and its corresponding buffer were added to 1 µg of Cas9D10A-GFP

and water was added to make a 50 µl reaction mixture. Below is the reaction component and corresponding volume

for a generic digestion 50 µl total volume reaction.

DNA (1 µg) 2 µl

Buffer 5 µl

Enzyme 1 µl

dH2O (nuclease free) 42 µl

50 µl total volume

The digestion was run at 37°C for 1 h. The resultant digest samples were run on an agarose gel.

Gel electrophoresis. 1.25 g of agarose was measured and mixed with 100 ml 1x TAE in a flask. The mixture was

microwaved for 1–3 minutes until the agarose was completely dissolved. The agarose solution was cooled to 20–

25°C. SYBR Safe gel stain was added to the solution. The 1.25% agarose solution was poured into a gel tray and

allowed to solidify. The agarose gel was placed into the electrophoresis unit and the unit was filled with 1x TAE.

The molecular weight ladder was loaded into the first lane of the gel. The digest samples were loaded into

subsequent lanes of the gel. The gel was run at 80 V for 1.5 h. The gel was visualized under UV light.

Sleeping Beauty transposon system. High fidelity Cas9 was cloned into the SfiI cloning sites in an empty SB

transposon vector with constitutive bi-directional promoter (pSbbi-GP). pSBbi-GP was a gift from Eric Kowarz

(Addgene plasmid # 60511). 106 MT-2 cells were nucleofected with 10.0 µg SB transposon plasmid (pSBbi-GP) and

5.00 µg SB transposase (SB100X). Cells used for nucleofection had >90% viability. 106 MT-2 cells were put into a

15 ml conical and washed once with PBS with the supernatant aspirated out. The cells were resuspended in 100 µl of

supplemented SE Nucleofector solution (2.25 ml SE Nucleofector solution + 0.5 ml supplement). Transposon and

transposase plasmid DNA was added to the 100 µl of cells in SE solution. 100 µl of the cell and DNA mixture was

transferred into a Nucleocuvette vessel and the vessel was placed in the 4-D Nucleofector X unit. The Jurkat CL-120

program was selected and run. After the run (~1 second), the Nucleocuvette vessel was removed and the contents

17

were transferred in 2 ml of 37°C RPMI 1640 + 2 mM Glutamine + 10% Fetal Bovine Serum (FBS) + 5 ml P/S.

18

2.3. Results

The goal of this project was to use CRISPR/Cas9-mediated genome editing to clone the novel heterozygous

CTLA4 mutation identified in our CVID patient into a Treg-like cell line to confirm this mutation as causal in the

autoimmune phenotype of the patient. There are many different methods to deliver and express Cas9 and the gRNA

in target cells that differ according to the goals of the experiment and ease of transfection of the target cells31-33.

However, the most common method uses transfection of the target cells with plasmids containing Cas9 and the

gRNA, as used by us in our workflow (Figure 3). Plasmids with Cas9 and the gRNA were constructed and

nucleofected into MT-2 cells. MT-2 cells are Treg-like cell line that has the phenotypic and functional characteristics

of human Treg cells36. This cell line was used as a model due to the fact that it constitutively expresses CTLA-4.

Since our aim was to model a specific point mutation as seen in our patient, we used a repair template in addition to

the gRNA and Cas9 in order to incorporate the mutation of interest into the cell through HDR. The patient mutation

is heterozygous and therefore the edited cell must only incorporate the mutation from the repair template into one

allele.

19

20

To increase the frequency of HDR, we used a Cas9 with only one active catalytic domain (Cas9 nickase)

that is only able to cut one strand, resulting in a nick rather than a DSB. Wild-type Cas9 has two nuclease domains,

HNH and RuvC, which cleave DNA by nicking the gRNA-complementary and noncomplementary strands,

respectively. We used a Cas9 with a D10A mutation in the RuvC domain (Figure 4). This Cas9 from Streptococcus

pyogenes requires a 5′-NGG PAM for target cleavage. We used a plasmid for Cas9 expression that contained Cas9

(D10A) nickase and GFP. The Cas9D10A-GFP plasmid was prepared as a maxiprep using a QIAprep Miniprep kit.

To introduce the monoallelic CTLA4P136A mutation into target cells, we selected a gRNA at the human

CTLA4 locus whose cleavage site is the least distance to the intended mutation with the lowest amount of off-target

sites (Figure 5). Although Cas9 generally cleaves its target reliably, off-target cleavage events can occur. Cas9

nuclease activity is guided by the gRNA and requires a PAM motif for cleavage of the target. Therefore, as long as

the Cas9 is able to recognize a guide sequence that is immediately upstream of a 5′-NGG PAM sequence, it will

cleave the DNA. Thus, if the guide sequence of the gRNA is nonspecific and is frequently present at sites outside of

the intended target site, off-site targeting and cleavage will occur31. To minimize off-target activity, an online guide

sequence design tool that computationally determines off-target cleavage activity was used to select a guide

sequence. The program estimates off-target activity of Cas9 at potential off-target sites by evaluating the frequency

of sequences within the genome with homology to the guide sequence or off-target sequences that vary by only a

21

few nucleotides as compared to the intended target.

A single-stranded oligonucleotide (ssODN) construct containing the intended mutation was used as a repair

template. For high HDR efficiency, ssODNs should contain sequences that flank the intended edit of at least 40 bp

that are homologous to the sequence of the target site, called homology arms. The ssODN sequence can be

homologous to the sense or antisense sequence of the target locus. We designed a 101-nt ssODN with 50 bp left and

right homology arms on either side of the intended C>G mutation (Figure 6a). The intended nucleotide in the

ssODN template is 4 bp away from the Cas9 cleavage site. The intended mutation is within the PAM of the gRNA

sequence and is expected to prevent re-cutting of Cas9 upon successful repair via the ssODN template (Figure 6b).

22

We first aimed to determine the efficiency of nucleofection by nucleofecting the Cas9D10A-GFP plasmid

into MT-2 cells and Jurkat cells. MT-2 cells are a human T-cell leukemia virus type 1 (HTLV-1) infected cell line

that has the phenotypic and functional characteristics of human Treg cells. A Treg-like cell line was used to study the

effect of the identified CTLA4 mutation given the role of CTLA-4 in Treg cell function. This cell line was used as a

model due to the difficulty of identifying and isolating CD4+FoxP3+ Treg cells. Jurkat cells were used as a control in

comparison of nucleofection efficiency for MT-2s based on their well-documented ability to undergo successful

nucleofection and their similarity in size and composition to MT-2s. 106 MT-2s and 106 Jurkat cells were

nucleofected with pmaxGFP to determine nucleofection efficiency. 106 MT-2s and 106 Jurkat cells were

nucleofected with the Cas9D10A-GFP plasmid. Control pmaxGFP plasmid-transfected MT-2s were used as a

reference for GFP expression. The efficiency of the Cas9D10A-GFP plasmid was determined by comparing live

versus dead cells and GFP expression of the positive control pmaxGFP plasmid versus the Cas9D10A-GFP plasmid.

48 hours post-nucleofection, flow cytometry was used to evaluate GFP expression and percentage of live

versus dead cells reported by staining of the cells with live/dead viability dye. Results showed very low levels of

live cells and GFP expression after nucleofection with the Cas9D10A-GFP plasmid. Previous Cas9 plasmid

preparation via maxiprep yielded low concentrations of plasmid DNA. The low quality Cas9 plasmid may have

lowered the efficiency of nucleofection since the quality and the concentration of DNA used for nucleofection is a

23

critical factor in the efficiency of gene transfer. The Cas9D10A-GFP plasmid was purified via maxiprep and the

DNA concentration was determined to be 495.5 ng/µl by UV spectrophotometry at 260 nm. The identity of the

plasmid was confirmed by gel electrophoresis.

Jurkat cells and MT-2 cells were nucleofected with the Cas9D10A-GFP plasmid after maxiprep

preparation. 30 hours post-nucleofection, the percentage of live versus dead cells and GFP expression was measured

by flow cytometry. Nucleofected cells showed 70% viability (Figure 7). Jurkat cells had a higher level of GFP

expression at 17% positive cells, whereas only 2% of MT-2s were positive for GFP expression. MT-2s showed a

nucleofection efficiency of 65% with the pmaxGFP control plasmid. A nucleofection efficiency rate of 65% is

consistent with expected efficiency rate as determined by optimization experiments. This suggests that MT-2s are

more resistant to nucleofection with larger plasmids than Jurkat cells. These results also demonstrate that the low

efficiency and high toxicity of the previous nucleofection was due to low quality plasmid DNA purification of the

Cas9D10A-GFP plasmid which was rectified upon re-purification of the plasmid.

The gRNA (MLM3636) and Cas9 (Cas9D10A-GFP) plasmids and ssODN were nucleofected into MT-2s

(Figure 8). The gRNA and Cas9 plasmids were nucleofected at a 1:1 molar ratio at a total of 2 µg of total DNA. The

24

ssODN template was nucleofected at an amount of 1 µl/500 ng. In order to obtain single-cell colonies, GFP+ cells

were single-cell sorted via flow cytometry into three 96-well plates. Growth was monitored every seven days for 21

days. A total of 288 cells were plated among which 0 colonies were recovered. These results suggest that MT-2s

may have not grown due to difficulty of the cell line to grow in single cell conditions.

The gRNA (MLM3636) and Cas9 (Cas9D10A-GFP) plasmids and ssODN were nucleofected into MT-2

cells. 9000 GFP+ live cells were bulk sorted via flow cytometry into 2 ml of conditioned RPMI medium (Figure 8).

24 hours post-sort, GFP+ MT-2s were viewed by microscopy. The cells were not visible under the microscope. Of

the 9000 cells that were positive for GFP, approximately 100 cells were visible. This result suggests an error during

the sorting process or a low tolerance of the cells to sorting process resulting in the loss of viable cells.

The gRNA (MLM3636) and Cas9 (Cas9D10A-GFP) plasmids and ssODN were again nucleofected into

MT-2 cells. 72 hours post-nucleofection, the cells were stained with live/dead viability dye and sorted for GFP

expression. 0 cells were positive for GFP, suggesting an error during the nucleofection procedure.

The nucleofection was repeated. The GFP+ cells were both bulk sorted and single-cell sorted. 181 GFP+

cells were bulk sorted into 1 ml of RPMI and 8 GFP+ cells were single-cell sorted into a 96-well plate. RPMI was

added every 3 days to the bulk sorted cells. The cells were expanded at 37°C with 5% CO2. These cells are still

currently growing out for sequencing.

25

We also constructed a plasmid that contained both the gRNA expression vector and Cas9 on the same

vector. Use of a plasmid that contains both the gRNA and Cas9 has the potential to allow higher throughput

introduction of pathogenic mutations into target cells by requiring construction and preparation of a single plasmid.

The 20-nt target sequence used in the MLM3636 plasmid was cloned into a vector backbone encoding a human U6

promoter-driven gRNA expression cassette and a CBh-driven Cas9-D10A (pSpCas9n(BB)-2A-GFP) (Figure 9). The

top strand oligo includes a guanine added to the 5′ end of the guide sequence not present in the target site in order to

increase transcription from the U6 promoter (Figure 9b, c).

The low efficiency of nucleofection observed in MT-2 cells is, at least partially, due to the large size of the

Cas9 plasmid. Stable expression of Cas9 by the target cell would negate the need for a Cas9 expression plasmid. The

26

use of transposable elements allows transgene integration of a gene of interest into a target cell. The Sleeping Beauty

(SB) transposable system was used to generate a MT-2 cell line that stably expresses Cas941. Plasmids that contain

the Sleeping Beauty transposon and transposase were nucleofected into MT-2 cells. The transposon plasmid has

Cas9, GFP, and puromycin, which is all stably integrated into genome of the cell. MT-2 cells were nucleofected

with pSBbi-GB and SB100X plasmids (Figure 10). 2 days after nucleofection, GFP+ cells were bulk sorted into

RPMI. 7 days post-sort, single cells were plated via limiting dilution to expand clonal populations.

27

28

3. Chapter 3: Discussion and Perspectives

3.1. Limitations

Widespread use of CRISPR-mediated gene editing for modeling disease-associated mutations is currently

limited by low HDR efficiency. This has prompted the development of various new strategies that all aim to increase

the efficiency of precise genome editing by use of CRISPR/Cas9 systems42-44. Although this work did not result in

an edited cell line with our patient mutation, it has furthered our search for an optimized protocol for modeling

mutations using CRISPR/Cas9.

Genome editing in cell lines via homologous-recombination-mediated CRISPR/Cas9 is inefficient due to

multiple variables. Delivery of DNA repair templates is difficult and methods for efficient transfection vary widely

between cell type. Difficult to transfect cell lines, like MT-2 cells, require the use of viral transduction or

nucleofection as transfection methods. Viral transduction has very high transfection efficiencies but requires that

DNA be introduced as viral vectors. Introduction of a repair template is not feasible using this approach.

Nucleofection can reach high transfection efficiencies and also allows us to deliver all necessary CRISPR/Cas9

system components.

Nucleofection of MT-2 cells has been successfully reported but is not well characterized. Nucleofection

programs are cell-type specific and apply specific high-voltage pulses dependent on cell type to facilitate transfer of

exogenous DNA into the nucleus. Thus, cytotoxicity and efficiency are dependent on optimization of the program

being used for the cell-type being nucleofected. MT-2 cells do not have a preset nucleofection program. Due to

similar structure and composition, we used a nucleofection program for Jurkat cells. Optimization data for a closely

related human T cell leukemia cell line called MT-4 reports a nucleofection efficiency of 60% for MT-4 cells. This

suggests that our program is optimal for MT-2 cells since we obtained 65.7% efficiency of nucleofection with the

control pmaxGFP plasmid. Thus, it is hypothesized that low GFP expression is likely due to reasons other than

nucleofection conditions.

The pmaxGFP plasmid is 3486 bp whereas the Cas9D10A-GFP plasmid is 9271 bp. Size is a factor for

uptake efficiency in nucleofection. Nuclear delivery of large plasmids is more difficult than small plasmids. This is

seen when using equivalent mass or molar concentrations of plasmid constructs of varying sizes. Alternative

methods for Cas9 expression in the target cell would negate the need for a Cas9 expression plasmid. Given the high

29

toxicity of nucleofection that leaves us with very few viable cells, using an approach to increase the percentage of

these viable cells that have successfully incorporated the foreign DNA is critical. The number of cells that we have

been able to recover after the nucleofection has been consistently low, despite the method we have used post-

nucleofection. This suggests that MT-2 cells are highly dependent on support from other cells to grow. Control

experiments we performed subjecting MT-2 cells to nucleofection without a plasmid present resulted in similar

results. This shows that the cell death observed is due to the nucleofection process itself and not to toxicity

associated with the plasmids or Cas9. Although sorting the cells is a stressor that the cells do not tolerate well,

selection of positive cells is necessary due to the low frequency of viable cells positive for plasmid incorporation.

Otherwise, the nucleofected cells would have to be single-cell sorted by limiting dilution. This would require a very

large number of cell populations to be expanded and sequenced, the vast majority of which would be negative for

plasmid incorporation. This process would be both expensive and time consuming with low throughput.

Transposable systems can be used for integration of a gene of interest into a target cell line. Stable gene

expression can be achieved using transposable elements (TEs) as a gene transfer system, negating the need for

plasmid-based gene expression. The Tc1/mariner-type Sleeping Beauty (SB) transposable system consists of a

transposase and a transposon that contains a gene-expression cassette for the gene that is to be transferred. The

transposase is an enzyme that catalyzes the movement of the transposon. The transposon plasmid contains a

promotor that directs transcription of the gene of interest. A second plasmid contains the gene for expression of the

transposase enzyme. The SB transposase is expressed and the enzyme binds to the inverted terminal repeats (ITR)

that flank the transposon and cuts the transposon out of the plasmid via an endonuclease reaction. The released

transposon can then bind a different DNA molecule with a TA sequence. The transposase creates a DSB in the DNA

which allows the transposon to integrate into the DNA. Since transgene integration occurs only at TA dinucleotides,

there is no insertion into promoters or first introns of actively transcribed genes as seen when using retro- and

lentiviral systems. The SB transposon vector can be designed for constitutive or inducible expression of the gene

expression41. The transposon plasmid used to generate our Cas9 stable MT-2 cell line has Cas9, GFP, and

puromycin, which is all stably integrated into genome of the cell. Although a large Cas9 expression plasmid is no

longer required for Cas9 expression in these cells, our use for this stable transgenic cell line is currently limited. The

gRNA plasmid must contain a selection marker of cells positive for the plasmid post-nucleofection. The MLM3636

gRNA plasmid used with the Cas9D10A-GFP plasmid did not contain a selection marker. GFP expression in our

30

nucleofected cells indicates successful uptake of the larger Cas9 plasmid. We thus used GFP expression for a marker

for uptake of both plasmids due to the smaller size of MLM3636 and the tendency that cells have to incorporate all

plasmids present together. Since the SB transposon vector contained a GFP expression cassette under a constitutive

promoter as part of the transgene, the MT-2 Cas9 stable cell line constitutively expresses GFP. Therefore, GFP

cannot be used to select for positive gRNA plasmid uptake in the stable cell line. Due to very low percentage of MT-

2 cells that are GFP+ after nucleofection, using a gRNA plasmid without a selection marker is not an option. Use of

a gRNA vector backbone with RFP was considered but was not an optimal option due to a difficult and expensive

cloning protocol. Although use of a gRNA-RFP vector may have allowed us to increase the efficiency of

nucleofection closer to the rates we observed with the control pmaxGFP plasmid, use of this method for cloning at a

higher-throughput is not favorable due to cost and difficulty.

3.2. Future Research

The Alt-R CRISPR/Cas9 system was recently developed by IDT as an alternative to traditional plasmid-

based delivery of CRISPR/Cas9 components. The Alt-R CRISPR/Cas9 system uses a Cas9 protein that is combined

with a crRNA:tracrRNA complex to form a ribonucleoprotein (RNP) complex, which is then delivered to the target

cells. The Alt-R CRISPR/Cas9 system uses a 2-oligo system of a 67-nt tracrRNA and a 36-nt crRNA. The lengths of

these RNAs were determined by systematic variation to provide the highest gene editing efficiency. The tracrRNA

can be fluorescently labeled for use in imaging or FACS sorting. The crRNA and tracrRNA are combined to form

the gRNA complex, which is then combined with Cas9 to form the RNP complex. The RNP and HDR template can

be delivered by lipofection or nucleofection. Although the components would still need to be delivered into MT-2s

by nucleofection, this method offers several potential advantages over the approach we have been using. This

system has shown higher on-target gene editing efficiency with less toxicity to the cells. By delivering our

CRISPR/Cas9 components as a RNP complex, this may reduce the toxicity observed in the cells post-nucleofection.

IDT also reports higher editing efficiency using the Alt-R system, which is still an issue for precise gene editing

even after optimizing delivery methods of the CRISPR/Cas9 system components.

The use of CRISPR/Cas9 for the insertion of precise genetic modifications is limited by the low efficiency

of HDR as compared to the high efficiency of NHEJ32-35. HDR is less frequent than NHEJ and only occurs during S

and G2 phase, whereas NHEJ occurs throughout the cell cycle. HDR does not occur sequentially but instead

concurrently with NHEJ. Inhibition of NHEJ improves the frequency of HDR and therefore methods to inhibit

31

components of the NHEJ pathway are being used to promote homologous recombination. Molecules involved in

NHEJ can be suppressed by gene silencing or inhibited by use of small molecule inhibitors. DNA ligase IV is a key

enzyme for NHEJ. Scr7 is a DNA ligase IV inhibitor and functions by targeting the DNA binding domain of DNA

ligase IV, reducing its affinity for DSBs and inhibiting its function. The use of Scr7 has been shown to increase the

efficiency of HDR-mediated genome editing in both mammalian cell lines and in mice up to 19-fold42, 43. NHEJ

repair can also suppressed by targeting DNA ligase IV using adenovirus 4 (Ad4) E1B55K and E4orf6 proteins,

which mediate the ubiquitination and proteasomal degradation of DNA ligase IV. Gene silencing of KU70, KU80,

or DNA ligase IV by short hairpin (sh)RNA have also been used for NHEJ suppression43. Inhibition of NHEJ can

induce apoptosis and therefore cell lines have varying sensitivity to treatment with NHEJ inhibitors. Manipulation of

cellular repair pathways can also affect the ability of the cell to respond to and repair damage at other non-target

sites, which may lead to tumor formation and, as a result, may not be feasible in therapeutic gene editing. However,

manipulation of cell cycle and cellular repair pathways can be used to increase the frequency of CRISPR/Cas9-

mediated precise gene modifications to optimize disease modeling in both cell lines and mouse models42-44.

Cas9-mediated HDR frequencies can also be increased by specific design of the repair template. Linearized

or double-stranded plasmid donors are often used as donor templates to incorporate large modifications or tags. For

small modifications or insertions, ssODN templates are more effective than plasmid donors. HDR events are more

frequent with repair templates with longer homology regions because the rate of recombination increases as the

length of homology arms increases. Homology arms of a ssODN can be as low as 40-nt but longer homology

regions have been reported to increase HDR efficiency. The length of ssODN donors can be optimized to increase

the incorporation rate of intended mutations. Longer ssODNs have more homology between the repair template and

the target site. However, longer ssODNs also have disadvantages in that they are more likely to be incorrectly

synthesized or have secondary structure45.

Recent research suggests that HDR frequency can be further improved by using donor ssDNA

complementary to the nontarget strand (—i.e., the sequence of the target strand). Donor–nontarget strand

complementarity increases the frequency of HDR events when using wild-type Cas9 or Cas9 variants (Cas9D10A or

Cas9H840A). It has also been recently demonstrated that homology-directed genome editing can be improved by

using asymmetric donor DNA as opposed to donor DNA symmetric around the break. Cas9 asymmetrically releases

DNA at the 3′ end of the cleaved DNA strand not complementary to the gRNA (nontarget strand) before complete

32

dissociation. Asymmetric donor DNA with 36 bp on the PAM-distal side of the Cas9 cut site and 91 bp on the

PAM-proximal side of the break complementary to the strand that is released first (nontarget strand) was found to

have the optimal design for annealing and is reported to increase the rate of HDR up to 60%45, 46.

The frequency of HDR can be improved using these methods, however, this does not address the accuracy

of HDR. Most HDR events do not result in the intended sequence change due to re-editing (Figure 11). Cas9 can re-

cut the edited locus, causing additional editing. The target locus can be re-cleaved until the guide sequence or PAM

is sufficiently modified by NHEJ that Cas9 is no longer able to recognize the target. HDR-mediated genome-editing

accuracy can be improved by blocking re-editing. Introducing CRISPR/Cas-blocking mutations in the guide

sequence or PAM blocks re-cutting by Cas9 and, as a result, improves HDR accuracy. CRISPR/Cas-blocking PAM

mutations are more efficient than gRNA sequence mutations. This is due to off-target activity by Cas9. Cas9 is able

to tolerate up to five mismatches in the gRNA sequence, which means that Cas9 may still cleave a DNA sequence

without complete guide sequence recognition45-47.

CRISPR/Cas9 editing is mostly biallelic and selective homozygous or heterozygous HDR-mediated gene

editing is not well characterized. Zygosity of the intended mutation can be controlled by designing the gRNA

sequence and donor template with specific cut-to-mutation distances. The full ssODN template is not always

incorporated during HDR. The probability that the mutation will be introduced decreases as the cut-to-mutation

distance increases. This relationship does not vary between loci. Thus, there is an inverse relationship between the

mutation incorporation rate and the distance from the cut site. Cut-to-mutation distance should be minimized for

homozygous mutation introduction. Biallelic editing can be achieved at higher rates by selecting a gRNA that

mediates a DSB at minimal distance (<10 bp) from the intended mutation site. In contrast, by selecting gRNA and

donor sequences with longer cut-to-mutation distances (2–26 bp), there is a lower likelihood that the mutation will

be incorporated and thus a higher frequency of heterozygous mutation incorporation. However, it is not always

feasible to select a gRNA sequence that will mediate a cut-to-mutation distance of <10 bp. Another method to

introduce monoallelic mutations in cases such as these is by using equimolar quantities of the donor template

containing the intended mutation and a donor template that is wild-type at the intended mutation site45, 47.

Homologous-recombination-mediated CRISPR/Cas9 gene editing offers great potential for modeling

human disease in cell lines and mouse models48-50. CRISPR-mediated gene editing can be used to model the

repression or activation of genes, disease-associated gene sequences, and epigenetic modifications involved in the

33

onset and progression of human disease. The use of the bacterial CRISPR/Cas9 system in editing and regulating

genomes has the capacity to change the way human disease is researched and treated. However, limitations prevent

its widespread use for precise gene editing. The future of CRISPR-mediated HDR depends on the development of

novel strategies to optimize gene editing. Given the variability that exists between cell lines and models, it is

important that different strategies be employed to develop an optimal protocol for the introduction of gene edits in

the model being used. Using several different approaches in parallel allows testing of these different approaches to

develop an efficient workflow in a timely manner. The combined use of various strategies to address the inefficiency

of HDR-mediated genome editing has high potential to lead to the development of a protocol that allows a high

throughput system for generating specific mutations.

34

4. Bibliography

1. Flajnik, M.F. & Kasahara, M. Origin and evolution of the adaptive immune system: genetic events and selective

pressures. Nat. Rev. Genet. 11, 47–59 (2010).

2. Bacchetta, R. & Notarangelo, L.D. Immunodeficiency with autoimmunity: beyond the paradox. Front. Immunol.

4, 77 (2013).

3. Maródi, L. & Notarangelo, L.D. Immunological and genetic bases of new primary immunodeficiencies. Nat.

Rev. Immunol. 7, 851–861 (2007).

4. Resnick, E.S. & Cunningham-Rundles, C. The many faces of the clinical picture of common variable immune

deficiency. Curr. Opin. Allergy Clin. Immunol. 12, 595–601 (2012).

5. Bogaert, D.J.A., Dullaers M., Lambrecht, B.N., et al. Genes associated with common variable

immunodeficiency: one diagnosis to rule them all? J. Med. Gen. 53, 575–590 (2016).

6. Maffucci, P., Filion, C.A., et al. Genetic diagnosis using whole exome sequencing in common variable

immunodeficiency. Front. Immunol. 7, 220 (2016).

7. Kuehn, H.S. et al. Immune dysregulation in human subjects with heterozygous germline mutations in CTLA4.

Science 345, 1623–1627 (2014).

8. Schubert, D. et al. Autosomal dominant immune dysregulation syndrome in humans with CTLA4 mutations.

Nat. Med. 20, 1410–1418 (2014).

9. Sakaguchi S. et al. Regulatory T cells and immune tolerance. Cell 133, 775–787 (2008).

10. Wing, K. et al. CTLA-4 control over Foxp3+ regulatory T cell function. Science 322, 271–275 (2008).

11. Wang, C.J. et al. CTLA-4 controls follicular helper T-cell differentiation by regulating the strength of CD28

engagement. PNAS 112, 524–529 (2015).

12. Sage, P.T., Paterson, A.M., Lovitch S.B. & Sharpe A.H. The coinhibitory receptor CTLA-4 controls B cell

responses by modulating T follicular helper, T follicular regulatory, and T regulatory cells. Immunity 41, 1026–

35

1039 (2014).

13. Brunet, J.F. et al. A new member of the immunoglobulin superfamily—CTLA-4. Nature. 328, 267–270 (1987).

14. Linsley, P.S. et al. CTLA-4 is a second receptor for the B cell activation antigen B7. J. Exp. Med. 174, 561–569

(1991).

15. Krummel, M.F. & Allison, J.P. CD28 and CTLA-4 have opposing effects on the response of T cells to

stimulation. J. Exp. Med. 182, 459–465 (1995).

16. Malissen, B. et al. Integrative biology of T cell activation. Nat. Immunol. 15, 790–797 (2014).

17. Schildberg, F.A. et al. Coinhibitory Pathways in the B7-CD28 Ligand-Receptor Family. Immunity 44, 955–972

(2016).

18. Zhang, X. et al. Crystal structure of the receptor-binding domain of human B7-2: Insights into organization and

signaling. PNAS 100, 2586–2591 (2003).

19. Pentcheva-Hoang, T. et al. B7-1 and B7-2 selectively recruit CTLA-4 and CD28 to the immunological synapse.

Immunity 21, 401–413 (2004).

20. Stamper, C.C et al. Crystal structure of the B7-1/CTLA-4 complex that inhibits human immune responses.

Nature 410, 608–611 (2001).

21. Yokosuka, T. et al. Spatiotemporal basis of CTLA-4 costimulatory molecule-mediated negative regulation of T

cell activation. Immunity 33, 326–339 (2010).

22. Schwartz, J.C. et al. Structural basis for co-stimulation by the human CTLA-4/B7-2 complex. Nature 410, 604–

608 (2001).

23. Darlington P.J., et al. Hierarchical regulation of CTLA-4 dimer-based lattice formation and its biological

relevance for T cell inactivation. J. Immunol. 175, 996–1004 (2005).

24. Qureshi, O.S. et al. Trans-endocytosis of CD80 and CD86: A molecular basis for the cell-extrinsic function of

CTLA-4. Science 332, 600–603 (2011).

36

25. Walker, L.S. & Sansom, D.M. The emerging role of CTLA4 as a cell-extrinsic regulator of T cell responses.

Nat. Rev. Immunol. 11, 852–863 (2011).

26. Kong, K.F. et al. Protein kinase C- η controls CTLA-4-mediated regulatory T cell function. Nat. Immunol. 15,

465–472 (2014).

27. Valk, Elke et al. CTLA-4 trafficking and surface expression. Trends Immunol. 29, 272–279 (2008).

28. Walker, L.S. & Sansom, D.M. Confusing signals: Recent progress in CTLA-4 biology. Trends Immunol. 36, 63–

70 (2015).

29. Marraffini, L.A. CRISPR-Cas immunity in prokaryotes. Nature 526, 55–61 (2015).

30. Jackson, S.A. et al. CRISPR-Cas: Adapting to change. Science 356, eaal5056 (2017).

31. Ran, F.A. et al. Genome engineering using the CRISPR-Cas9 system. Nat. Protoc. 8, 2281–2308 (2013).

32. Sander, J.D. & Joung, J.K. CRISPR-Cas systems for editing, regulating and targeting genomes. Nat. Biotechnol.

32, 347–355 (2014).

33. Mali, P., Esvelt K.M. & Church, G.M. Cas9 as a versatile tool for engineering biology. Nat. Met. 10, 957–963

(2013).

34. Mali, P. et al. RNA-guided human genome engineering via Cas9. Science 339, 823–826 (2013).

35. Jiang, W., Bikard, D., Cox, D., Zhang, F. & Marraffini, L.A. RNA-guided editing of bacterial genomes using

CRISPR-Cas systems. Nat Biotechnol. 31, 233–239 (2013).

36. Hamano, R. et al. Characterization of MT-2 cells as a human regulatory T cell-like cell line. Cellular &

Molecular Immunol. 12, 780–782 (2015).

37. Lieber, M.R. The mechanism of double-strand DNA break repair by the nonhomologous DNA end-joining

pathway. Annu. Rev. Biochem. 79, 181–211 (2010).

38. San Filippo, J., Sung, P. & Klein, H. Mechanism of eukaryotic homologous recombination. Annu. Rev.

37

Biochem. 77, 229–257 (2008).

39. Davis, L. & Maizels, N. Homology-directed repair of DNA nicks via pathways distinct from canonical double-

strand break repair. PNAS 111, E924–E932 (2014).

40. Symington, L. S. & Gautier, J. Double-strand break end resection and repair pathway choice. Annu. Rev. Genet.

45, 247–271 (2011).

41. Kowarz, E., Löscher, D., & Marschalek, R. Optimized Sleeping Beauty transposons rapidly generate stable

transgenic cell lines. Biotechnol. J. 10, 647–653 (2015).

42. Maruyama, T. et al. Increasing the efficiency of precise genome editing with CRISPR-Cas9 by inhibition of

nonhomologous end joining. Nat. Biotechnol. 33, 538–542 (2015).

43. Chu, V.T. et al. Increasing the efficiency of homology-directed repair for CRISPR-Cas9-induced precise gene

editing in mammalian cells. Nat. Biotechnol. 33, 543–548 (2015).

44. Lin, S., Staahl, B.T., Alla, R.K. & Doudna, J.A. Enhanced homology-directed human genome engineering by

controlled timing of CRISPR/Cas9 delivery. elife 3, e04766 (2015).

45. Kwart, D., Paquet, D., Teo, S. & Tessier-Lavigne, M. Precise and efficient scarless genome editing in stem cells

using CORRECT. Nat. Protoc. 12, 329–354 (2017).

46. Richardson, C.D., Ray, G.J., DeWitt, M.A., Curie, G.L. & Corn, J.E. Enhancing homology-directed genome

editing by catalytically active and inactive CRISPR-Cas9 using asymmetric donor DNA. Nat. Biotechnol. 34,

339–344 (2016).

47. Paquet, D. et al. Efficient introduction of specific homozygous and heterozygous mutations using

CRISPR/Cas9. Nature 533, 125–129 (2016).

48. Li, K. et al. Optimization of genome engineering approaches with the CRISPR/Cas9 system. PLoS ONE 9,

e105779 (2014).

49. Dow, L.E. Modeling disease in vivo with CRISPR/Cas9. Trends Mol. Med. 21, 609–621 (2015).

38

50. Hsu, P.D., Lander, E.S. & Zhang, F. Development and applications of CRISPR-Cas9 for genome engineering.

Cell 157, 1262–1278 (2014).